学 位 論 文 Studies on the activation mechanism of Hedgehog signaling in fish 魚類におけるヘッジホッグシグナル伝達経路の 活性化機構の解析 平成25年7月博士(理学)申請 東京大学大学院理学系研究科 生物科学専攻 山 元 孝 佳 ( )
学 位 論 文
Studies on the activation mechanism of
Hedgehog signaling in fish
魚類におけるヘッジホッグシグナル伝達経路の
活性化機構の解析
平成25年7月博士(理学)申請
東京大学大学院理学系研究科
生物科学専攻
山 元 孝 佳
(
)
2
Contents
Contents ............................................................................................................................ 2
Abbreviations ................................................................................................................... 7
Abstract ............................................................................................................................. 9
Introduction ..................................................................................................................... 11
Results ............................................................................................................................ 18
Generation of maternal-zygotic aA90/dhc2 mutants .................................................. 18
Patterning of the spinal cord in MZdhc2 mutants ....................................................... 22
Lower Hh pathway activation in mutant cells ............................................................ 25
Patched1 localizes to cilia in medaka fish .................................................................. 26
MZdhc2 cells are less sensitive to Shh ....................................................................... 27
Fused forms a positive-feedback loop in fish ............................................................. 29
Discussion ....................................................................................................................... 32
A possible role of cilia in Hh gradient formation ........................................................ 32
Importance of cilia in Hedgehog signaling ................................................................. 34
3
Significance of teleost-specific augmentation of Hh signaling mediated by Fused ... 36
Conclusions .................................................................................................................... 39
Materials and methods .................................................................................................... 40
Fish strains .................................................................................................................. 40
Genotyping of medaka aA90/dhc2 mutant .................................................................. 41
Antibody generation .................................................................................................... 41
Whole mount in situ hybridization .............................................................................. 42
mRNA overexpression ................................................................................................ 43
Microinjection and Cell transplantation ...................................................................... 43
Histology ..................................................................................................................... 45
Immunohistochemistry ................................................................................................ 46
Chemical treatment ..................................................................................................... 47
RT-PCR ....................................................................................................................... 47
Scanning electron microscope .................................................................................... 48
Figures ............................................................................................................................ 49
4
Figure 1. The French flag model providing a positional information by a morphogen
concentration gradient. ................................................................................................ 50
Figure 2. Hedgehog signal transduction pathway. ...................................................... 51
Figure 3. Schematic view of neural tube patterning by Shh concentration gradient. . 52
Figure 4. The formation and maintenance of cilia mediated by the intraflagellar
transport. ..................................................................................................................... 53
Figure 5. Morphological phenotypes of aA90/dhc2 mutants. ..................................... 55
Figure 6. The medaka aA90/dhc2 lacks essential domains of the dhc2 gene. ............ 57
Figure 7. Generation of Maternal-Zygotic dhc2 mutant (MZdhc2). ........................... 59
Figure 8. Cilia are shortened in MZdhc2. ................................................................... 61
Figure 9. Neural tube patterning in MZdhc2 mutants. ................................................ 63
Figure 10. Gross patterning of neural tube and somite in MZdhc2 mutant. ............... 65
Figure 11. Dose-dependent effects of cyclopamine treatment on the expression of Hh
target genes.................................................................................................................. 67
Figure 12. nkx2.2 and olig2 expression at three anterior-posterior axis levels. .......... 68
5
Figure 13. A schematic drawing explaining the similarities and differences in ciliary
and neural tube phenotypes between fish and mouse dhc2/dnchc2 mutants. ............. 70
Figure 14. Hh signaling activity is partially defective in MZdhc2 mutants. .............. 71
Figure 15. MZdhc2 is sensitive to dnPKA. ................................................................. 72
Figure 16. Ptch1 is localized to the cilia in medaka fish. ............................................ 74
Figure 17. Ectopic olig2 expression of WT cells in the dorsal region of MZdhc2
neural tube. .................................................................................................................. 76
Figure 18. fused is required for Hh signaling in medaka fish. .................................... 79
Figure 19. fused expression pattern in medaka and zebrafish, and fused augments Hh
signaling in medaka. ................................................................................................... 80
Figure 20. Proposed model of the distinct features of Hh signal transduction in insect,
fish and mammal. ........................................................................................................ 82
Tables .............................................................................................................................. 83
Table 1. Defects in heart asymmetry in dhc2 mutant embryos and morphants .......... 84
Table 2. Primers used in this study ............................................................................. 85
6
Table 3. Accession numbers used to create the phylogenetic trees depicted in Fig. 16A.
..................................................................................................................................... 86
Table 4. Number of samples to examine Hh activity with the graded series of
cyclopamine treatment depicted in Fig. 14A-B. ......................................................... 87
References ...................................................................................................................... 88
Acknowledgements ........................................................................................................ 92
7
Abbreviations
dhc2 cytoplasmic dynein heavy chain 2
dhh desert hedgehog
dpf days post-fertilization
ENU N-ethyl-N-nitrosourea
fu fused
GFP green fluorescent protein
HC heavy chain
Hh hedgehog
IC intermediate chain
IFT intraflagellar transport
ihh indian hedgehog
KV Kupffer's vesicle
LNT lateral neural tube
Mdhc2 maternal mutant of dhc2
8
MeOH methanol
MO morpholino antisense oligonucleotide
MZdhc2 maternal-zygotic mutant of dhc2
ORF open reading frame
PFA paraformaldehyde
Ptch1 Patched 1
shh sonic hedgehog
Smo Smoothened
VM ventral midline
WT wild type
Zdhc2 zygotic mutant of dhc2
9
Abstract
Primary cilia are essential for Hedgehog (Hh) signal transduction in vertebrates.
Although the core components of the Hh pathway are highly conserved, the dependency
on cilia in Hh signaling is considered to be lower in fish than in mice, suggesting the
presence of species-specific mechanisms for Hh signal transduction.
To precisely understand the role of cilia in Hh signaling in fish and explore the
evolution of Hh signaling, I have generated a maternal-zygotic medaka (Oryzias latipes)
mutant that lacks cytoplasmic dynein heavy chain 2 (dhc2; MZdhc2), a component
required for retrograde intraflagellar transport. I found that MZdhc2 exhibited the
shortened cilia and partial defects in Hh signaling, although the Hh defects were milder
than zebrafish mutants which completely lack cilia. This result suggests that Hh activity
in fish depends on the length of cilium. However, the activity of Hh signaling in
MZdhc2 appeared to be higher than that in mouse dhc2 mutants (also called Dnchc2),
suggesting a lower requirement for cilia in Hh signaling in fish. I have revealed that the
receptor Ptch1 is exclusively localized on the cilium in fish as in mammals. Subsequent
10
analyses revealed that Fused, an essential mediator for Hh signaling in Drosophila and
fish but not in mammals, augments the activity of Hh signaling in fish as a
transcriptional target of Hh signaling. The finding of this fish-specific augmentation
provides a novel insight into the evolution of Hh signaling.
11
Introduction
Sexually reproducing multicellular organisms are developed from a single cell, a
fertilized egg. Through numerous times of cell divisions, the egg gives rise to hundreds
of different cell types, such as neurons, eyes, germ cells and muscles. These different
tissues and organs do not exhibit random distribution but are organized with a
remarkable reproducibility during development; eyes are always located on the head,
not on our legs or arms, while the brain is placed inside the skull. How are these tissues
and organs reproducibly orchestrated?
Morphogen gradient formation is a key concept for understanding these
organizations. Morphogen is a diffusible molecule secreted into the extracellular space
from its source and makes a concentration gradient. Multiple cell types are
differentiated depending on its concentration, as in a model, so called "French flag
model" (Rogers and Schier, 2011) (Fig. 1).
This concept of morphogen gradient has been used for understanding the
regeneration of hydra and planarian flatworms since the 1700s. When these animals are
12
cut into two halves, the head half regenerates a tail, while the tail half regenerates a head
half. This suggests that a kind of "polarity" is present along the body axis. In 1924,
Spemann and Mangold discovered that transplantation of Spemann organizer, a cluster
of dorsal cells in an amphibian gastrula embryo into the ventral region of a host gastrula
induces a secondary axis, suggesting that inducing signals are released from the explant
(Spemann and Mangold, 2001). In 1952, Turing proposed that chemical substances,
called morphogens, can provide positional information for cells by making a
concentration gradient (Turing, 1952).
Hedgehog (Hh) is one of the important morphogens in animal development,
which is evolutionarily conserved from fly to human. Hh protein functions by binding
to cell-surface receptor Patched, which serves as an inhibitor of Smoothened (Smo), a
downstream membrane-bound mediator of Hh signaling. When Hh ligand binds to the
receptor, the shape of Patched protein is altered so that it no longer inhibits the activity
of Smo, thereby leading to the activation of Gli/Ci (short for Cubitus interruptus), a zinc
finger containing transcriptional factor (Fig. 2).
13
hh was first identified as a segment polarity gene in Drosophila
(Nusslein-Volhard and Wieschaus, 1980). Vertebrates have three homologues of the
gene: sonic hedgehog (shh), desert hedgehog (dhh) and indian hedgehog (ihh) (Pathi et
al., 2001). The expression patterns of these three paralogues are largely not overlapped.
dhh is expressed in the Sertoli cells of the testes, essential for spermatogenesis and ihh
is in the gut and cartilage. shh is expressed in many tissues, has the most variety of
functions among the three homologues, and is essential for various aspects of
embryogenesis including patterning events of the neural tube and limb (Huangfu and
Anderson, 2006; McMahon et al., 2003). For example, in the neural tube, the sonic
hedgehog (Shh) ligand forms a dorso-ventral (DV) gradient with the highest
concentration ventrally, and specifies cell fates in a concentration-dependent manner
(Dessaud et al., 2008). Thus, the expression of cell-type specific genes serves as a
readout of Hh activity and delineates domains in the ventral neural tube (Fig. 3). The
mechanism of Hh-signal transduction has been the target of intense studies but remains
only partially understood.
14
One of the striking features of Hh signaling is that the primary cilium, a
microtubule-based, immotile cellular protrusion, is essential in vertebrates but not in
Drosophila (Wilson and Chuang, 2010). A requirement for the cilium in this pathway
was first identified by genetic screening in mice for ciliary mutants exhibiting
phenotypes similar to those of Hh-pathway mutants (Huangfu et al., 2003). However,
subsequent genetic and molecular analyses demonstrated that cilium-dependency and
the mediators of Hh signaling varies between fish and mammals, raising a question
about conservation and evolution of the mechanism of Hh-signal transduction (Huang
and Schier, 2009).
The formation and maintenance of cilia depend on the conserved process of
intraflagellar transport (IFT) (Goetz and Anderson, 2010). Ciliary proteins are
transported along the ciliary axoneme by IFT machinery, driven by kinesin-based
anterograde and dynein-powered retrograde transport (Fig. 4).
In general, cilia are classified into two types; one is a motile cilium and the
other is an immotile one, so called primary cilium. Motile cilia are present in some
specific tissues such as the epithelium of the ventricle, trachea and oviduct. Defective
15
cilia have been implicated in numerous diseases termed ciliopathies, including
hydrocephalus, bronchitis, infertility and situs inversus. Primary cilia are ubiquitously
found on vertebrates cells with a few exceptions of myeloid and lymphoid (Wheatley,
1995). These cilia were first described in 1898 (Zimmermann, 1898), but their functions
during animal development and physiology are only beginning to be unveiled
(Eggenschwiler and Anderson, 2007).
Recently, the cilium was reported to resemble the nucleus in terms of the
protein localizations and transport machinery. The nuclear import machinery including
GTP-bound Ran and importin-β2 are also involved in ciliary import (Dishinger et al.,
2010) and some nuclear pore complex proteins are located at the base of cilia and
possibly function as a diffusion barrier (Kee et al., 2012). These results imply that cilia
could have some essential roles in a transcriptional regulation.
Importantly, mammalian primary cilia have been recently shown to mediate
transduction of Hedgehog (Hh) signals (Huangfu et al., 2003; Wilson and Chuang,
2010). Further analysis provided that several key components of Hh pathway are
enriched in cilia, including Ptch1, Smo and Gli transcription factors (Goetz and
16
Anderson, 2010). In the absence of Ift88, a component of anterograde IFT machinery,
both mouse and zebrafish embryos lack cilia and exhibit a severe reduction in Hh
signaling. However, the phenotype is milder in zebrafish. In the neural tube, most of the
Hh target genes are not expressed in mouse mutants, while the expression of
low-threshold genes remains and expands in zebrafish (Huang and Schier, 2009;
Huangfu et al., 2003). These results suggest that cilium is required for Hh signaling also
in fish, but the dependency on cilia is lower than that in mammals. However, it was still
unclear how much Hh signaling in fish depends on cilia and what is the underlying
mechanism for that difference.
Furthermore, Fused (Fu), a putative serine-threonine kinase, first identified as
an essential mediator of Hh signaling in Drosophila, turned out to be not required for
mammals, but it is indispensable for zebrafish (Chen et al., 2005; Merchant et al., 2005;
Wilson et al., 2009; Wolff et al., 2003). These facts suggest that the pathway in zebrafish
is more similar to that in Drosophila or placed in between Drosophila and mammals,
making fish a unique model with which to investigate the transition state from an
ancestral to a modern type of Hh signaling.
17
To further address the difference and the evolution of Hh signaling among
species, I have generated a maternal-zygotic (MZ) medaka mutant that lacks
cytoplasmic dynein heavy chain 2 (dhc2), an essential component of retrograde IFT, and
compared the neural phenotypes of medaka and mouse dhc2 mutants (also called
Dnchc2, especially in mouse). I have generalized that the requirement for cilia in Hh
signaling is lower in fish than in mammals. Additionally, the receptor Ptch1 is localized
to cilia in fish as in mammals. Subsequent analyses revealed that the difference in the
requirement for cilia in Hh signaling across vertebrates can be interpreted by differential
regulation and function of Fu.
18
Results
Generation of maternal-zygotic aA90/dhc2 mutants
The medaka aA90 (Zdhc2) mutant, isolated in an ENU-induced mutagenesis screening
(Yokoi et al., 2007), is a recessive lethal mutant showing defects in left-right (L/R) axis
determination (Fig. 5A-B; Table 1). L/R asymmetry is established by directional flow of
extra-embryonic fluid surrounding the node (Kupffer's vesicle in fish) by cilia (Nonaka
et al., 1998). To identify the defective gene in the aA90 mutant, Dr. Tadashi Ishiguro (a
previous undergraduate student) carried out positional cloning and narrowed down the
aA90 locus to a 250 kb region in linkage group 13, which harbors one open reading
frame, cytoplasmic dynein heavy chain 2 (dhc2), an IFT retrograde component (Fig. 4,
6A). He found that aA90 has a 37.7 kb deletion in the dhc2 locus including the start
codon, the heavy chain (HC)-HC and the HC-Intermediate chain interaction domain,
and the AAA ATPase domain (Fig. 6A, C). Database searches demonstrated that the
dhc2 gene exists as a single copy within the medaka genome. I injected antisense
morpholinos (MO) against the dhc2 gene into wild-type embryos significantly
19
phenocopied aA90, which led me to conclude that dhc2 is the gene deficient in the aA90
mutant (Table 1). I named aA90 mutant as dhc2 mutant in the following description.
Probably due to the maternal contribution of dhc2-gene products, the
phenotype of dhc2 mutants was mild. For example, only one-fourth of the dhc2
homozygous mutants showed situs inversus (Table 1). To completely eliminate dhc2
products, I have generated maternal-zygotic dhc2 (MZdhc2) mutants using the
germline-replacement technique (Ciruna et al., 2002; Shimada and Takeda, 2008) with
the following modifications (Fig. 7). For making a maternal-zygotic medaka mutant, it
is known to use the interspecific hybrid sterility of Japanese (Kaga) and Chinese
(Hainan) medaka as hosts for transplantation of germ cells from homozygous donors
followed by sterility check of the hosts after sexual maturation (Shimada and Takeda,
2008). The hybrid, however, does not produce many eggs. In zebrafish, the knockdown
technique using morpholino against an essential gene, dead end, for the primordial germ
cell (PGC) has been used for making a maternal zygotic mutant (Ciruna et al., 2002). I
decided to apply the zebrafish knockdown method to medaka. As a host, I used
olvas-GFP transgenic medaka, whose oocytes were labeled with green fluorescent
20
proteins using the regulatory region of the medaka vasa gene (olvas, named after
Oryzias latipes vasa) (Tanaka et al., 2001), so that I easily check whether they have
germ cells or not. To label donor cells, I injected the rhodamine-dextran (10 kDa;
Molecular Probes, D1816), instead of "fixable" rhodamine dextran (10 kDa; Molecular
Probes, D1817) which was frequently used in medaka transplantation experiments,
considering the toxicity for the early embryogenesis in medaka. Host embryos were
injected at one-cell stage with 300 μM of a morpholino antisense oligonucleotide
directed against dead end mRNA and checked the loss of their germ cells at 2 dpf by the
loss of GFP fluorescence. Donor embryos were genotyped to identify homozygous
embryos. The other procedures were done according to the previous report (Shimada
and Takeda, 2008). Crosses of females with mutant germ cells and heterozygous males
(dhc2/+) generated 50% homozygous mutants that lacked both maternal and zygotic
products of dhc2 (MZdhc2) and 50% heterozygous mutant embryos that lacked only the
maternal dhc2 contribution (Mdhc2). As expected, hosts produced many eggs like wild
type adults.
21
The complete loss of dhc2 activity increased the frequency of situs inversus to
52.8% (Table 1, Fig. 5A-D) as well as enlarged ventricles and expanded nephric duct
(Fig. 5E-H, M-P). Moreover, the typical phenotypes of defective Hh-signaling, severe
ventral curvature and U-shaped somites instead of chevron-shaped ones, were observed
in MZdhc2 mutants, but not in zygotic (Zdhc2) or Mdhc2 mutants (Fig. 5I-L, Q-T),
indicating reduced levels of Hh signaling. Importantly, the morphology of cilia was
dramatically shortened in MZdhc2 as demonstrated by scanning electron microscopy
(SEM) (Fig. 8A). To expose the ventricular surface area of neural tubes, I exteriorized
this area with forceps, prior to fixation (Fig. 8B) and found that cilia on the surface of
non-floor plate (FP) neuroepithelial cells (LNT, lateral neural tube) and longer ones on
the FP cells (VM, ventral midline) were much shorter and bloated in MZdhc2 than their
wild-type counter parts (Fig. 8A). In the Kupffer’s vesicle and somites, cilia were also
shortened in MZdhc2, as compared with those in WT, Mdhc2 and Zdhc2 (Fig. 8C, data
not shown). The number and morphology of cilia in Zdhc2 mutants appeared normal at
least until the segmentation stages, but subtle defects in function or lately overt defects
could account for their milder phenotypes (Table 1, data not shown). The ciliary
22
phenotypes in MZdhc2 mutants are nearly identical to those in mouse dhc2/Dnchc2
mutant (Huangfu and Anderson, 2005; May et al., 2005), and thus the analysis of the Hh
activity in MZdhc2 mutants enabled us to examine differences and distinct mechanisms
between fish and mouse in the requirement for cilia in Hh signaling.
Patterning of the spinal cord in MZdhc2 mutants
As described in Introduction (Fig. 3), depending on the Hh gradient, the vertebrate
neural tube exhibits position-specific gene expression along the dorso-ventral axis;
roughly from ventral to dorsal, foxa2 in the FP, nkx2.2 in p3 neuron precursors, olig2 in
motor neuron precursors (pMN), nkx6.1/6.2 in p3/pMN/p2 progenitors, and pax6, pax3,
dbx1 and dbx2 in dorsally located neuron precursors and their expressions are mutually
exclusive underlined by their repressive interactions (Fig. 3) (Dessaud et al., 2010;
Jeong and McMahon, 2005). Shh is known to induce the expression of the ventral genes
(foxa2, nkx2.2, olig2, nkx6.1and nkx6.2), while suppressing the dorsal genes (pax6, pax3,
dbx1 and dbx2) (Balaskas et al., 2012; Dessaud et al., 2010; Jeong and McMahon, 2005).
I first confirmed that shh was normally expressed in the medial FP (MFP) and
23
underlying notochord of MZdhc2 mutants (Fig. 9A, 10J), suggesting that defects
observed in mutants are mainly ascribed to signal transduction defects.
In MZdhc2 mutants, foxa2 and nkx2.2 were expressed (Fig. 9A, 10H-I), and
ventral intermediate genes, olig2, nkx6.1 and nkx6.2 were dorsally expanded, whereas
this dorsal expansion was not observed in Zdhc2 (Fig. 9A, 10E-G).
The expression of these ventral genes suggests that the Hh pathway is activated
in cells with severely shortened cilia and even reaches the high levels of activation, not
missing the expression of the most ventral side genes. Additionally, like zebrafish, the
most ventral gene foxa2 expression in the medial FP is Hh-independent in medaka
embryos (Fig. 11B), and thus I will use nkx2.2 expression as a marker of the high level
of Hh activation. Also, it is worth noting that the most ventral region appeared to be
missing in MZdhc2 embryos, because the expression domains of nkx2.2, which are
normally separated by the negative medial FP cells, frequently merged in the medial
region (Fig. 9A). However, due to the lack of a specific marker for this region, I was
unable to determine the cell type specifically defective in MZdhc2 embryos.
24
The expansion of lower-threshold gene expression (olig2, nkx6.1 and nkx6.2)
also suggests that the area of low Hh activation abnormally expanded dorsally in the
mutant neural tube, probably because shortened cilia in MZdhc2 could not retain Hh
ligand, leading to dorsally expanded distribution of the ligand (further analysis and
discussion are described in the result of "MZdhc2 cells are less sensitive to Shh" and
discussion section). Dorsal expansion of olig2 expression in MZdhc2 was also observed
at three different anterior-posterior axis levels (Fig. 12). This was further supported by
dorsally retracted expression of pax6, pax3, dbx1 and dbx2, observed in MZdhc2
mutants (Fig. 9A, 10A-D), reflecting the repressive interactions of these genes.
In Dnchc2-mutant mice, nkx2.2 expression was reported to be lost, but olig2
was expanded (Huangfu and Anderson, 2005; May et al., 2005). Thus, there are
similarities and differences in the neural tube phenotypes between fish and mouse dhc2
mutants (Fig. 13), both of which I addressed in the following experiments.
25
Lower Hh pathway activation in mutant cells
To examine the activation level of Hh pathway in mutant cells, I treated MZdhc2
embryos with various concentrations of cyclopamine, a potent antagonist of
Smoothened (Smo). Intriguingly, in the MZdhc2 group, the percentage of
nkx2.2-positive embryos started to decrease at a cyclopamine concentration as low as
0.25 μM, and went down below 50% at 0.5 to 1 μM, while at such low concentrations,
100% of embryos maintained nkx2.2 expression in the wild-type and Mdhc2 groups
(Fig. 14A, C; Table 4). These results suggest that the activity of Hh signaling in mutant
cells is compromised at the level or upstream of Smo, but still high enough to express
the ventral-most marker, nkx2.2.
Zebrafish mutants with complete lack of cilia were reported to be insensitive to
cyclopamine and dominant-negative (dn) PKA (Ben et al., 2011; Huang and Schier,
2009), which induces ectopic Hh-pathway activation downstream of Smo. By contrast,
MZdhc2 is sensitive to cyclopamine (Fig. 14A, C) and dnPKA (Fig. 15), probably
because they have shortened but certain cilia. This discrepancy can be explained by the
fact that cilia are required for both Smo and PKA activity in fish. This is why the
26
MZdhc2 mutant is sensitive, and the mutants with a complete loss of cilia are not
sensitive, to cyclopamine and dnPKA.
Patched1 localizes to cilia in medaka fish
In murine cells, Hh receptor Patched1 (Ptch1) was reported to localize the primary
cilium at least in cultured cells and paraxial mesoderm cells (Ocbina et al., 2011;
Rohatgi et al., 2007), whereas it is not the case in Drosophila, which does not require
cilia for the reception of Hh (Wilson and Chuang, 2010). However, the receptor
localization was unknown in fish. Two homologues (patched1 and patched2) of
Drosophila patched were isolated in fish. To distinguish these two paralogues, I made
the phylogenic tree of these patched genes (Fig. 16A). As a result, medaka Ptch1 is the
homologue of mammalian Ptch1, which is a receptor of Shh. For further confirmation, I
knocked down Ptch1 in medaka and analyzed the Hh activity. In the morphants, the
number of Engrailed (a Hh target gene) expressing cells in somites was increased (Fig.
16B), representing the increased Hh activation due to the lack of the receptor, similar to
zebrafish morphant (Wolff et al., 2003). These results indicate that Ptch1 is the receptor
27
of Hh in medaka. To analyze the receptor localization in medaka, I generated an
antibody against the extracellular domain of medaka Ptch1 (Fig. 16C), and examined
the distribution of Ptch1 in wild-type and MZdhc2 neural-tube cells. Firstly, the
specificity of the antibody was confirmed by knockdown and overexpression
experiments (Fig. 16D, E). As shown in Figure 16D, in WT, Ptch1 was localized to the
cilia of neuroepithelial cells which were exteriorized with forceps before fixing (Fig.
8B). Importantly, Ptch1 was still localized to severely shortened cilia in MZdhc2 (Fig.
16D). These results indicate that the cilium is the site for Hh receptor Ptch1 localization
in medaka.
MZdhc2 cells are less sensitive to Shh
Although the activation level of Hh signaling is still sufficient to induce all target genes
in MZdhc2 cells, the amount of Ptch1 in severely shortened cilia is likely to be
decreased. This could explain the higher sensitivity to cyclopamine in MZdhc2 than that
in WT in the above experiment (Fig. 14A, C). In other words, MZdhc2 cells could be
less sensitive to Shh. Although the transplantation of mutant cells into wild-type neural
28
tubes was a straight way to test this idea, I thought that it was hard for me to detect the
loss of target gene expression in a single cell embedded in cells positive for target gene
expression. Thus, I transplanted wild-type cells into MZdhc2 blastula or Mdhc2
(control), and examined olig2 expression when donor cells were localized in host neural
tubes (Fig. 17A). I distinguished MZdhc2 from Mdhc2 embryos by the eye phenotype at
16-somite stage when transplanted embryos were fixed for the analysis (Fig. 17B). The
defect in eye formation can be explained by the fact that the rods and cones of the retina
consist of highly modified cilia. Remarkably, olig2-positive WT cells were frequently
found in the region more dorsal to the host olig2-expression domain in MZdhc2
embryos (Fig. 17C, WT to MZdhc2 (right panel), arrowhead; n=9/10), while no such
ectopic expression was detected in control transplants (Fig. 17C, WT to Mdhc2 (left
panel), arrowhead; n=15/15). These results demonstrate that Hh-activation of MZdhc2
cells is lower than that in WT cells, even if they are exposed to the same concentration
of Hh-ligand. Additionally the ectopic expression suggests that the distribution of Hh
ligand is dorsally expanded in MZdhc2, due to the decreased number of the receptor
available on the cell surface.
29
Fused forms a positive-feedback loop in fish
The presence of nkx2.2 expression is unique in MZdhc2, considering that mouse
Dnchc2 mutants lose nkx2.2 expression (Huangfu and Anderson, 2005; May et al.,
2005). The same tendency was observed in the expression analysis of Hh target genes of
ift88 mutants in zebrafish and mouse that completely lack cilia; only zebrafish mutants
maintain the expression of intermediate genes like olig2 (Huang and Schier, 2009;
Huangfu et al., 2003), implying that the activation of Hh signal is enhanced in fish. To
explore a teleost-specific mechanism, I focused on fused (fu), an intracellular mediator
of Hh signaling downstream of Smo in Drosophila, which has evolved divergent roles
in the vertebrate lineage: one for Hh signaling and the other for ciliary motility.
Interestingly, murine Fu is not involved in Hh signaling and specifically participates in
the motility of cilia, whereas it is required for both in zebrafish (Wilson et al., 2009;
Wolff et al., 2003). I first tested whether fu is essential for Hh signaling in medaka by
injecting fu MO (600 μM) targeted to the splicing site (Fig. 18A) and observed the loss
of nkx2.2 expression (Fig. 18B; n=14/15). Additionally, morphants injected together
30
with fu mRNA rescued nkx2.2 expression (Fig. 18C; n=14/14) and injection of fu
mRNA into WT embryos elevated Hh activity as indicated by the expansion of the
ventral intermediate genes, olig2 and nkx6.1 (Fig. 18D; n=9/14, 9/9, respectively). I
then knocked down fu in MZdhc2 mutants to see if the remaining expression of Hh
target genes in those mutants also depends on Fu. However, under this experimental
conditions, most of the MZdhc2 mutants injected with fu MO (600 μM) died probably
due to a requirement of Fu in earlier development (Xia et al., 2010), and I therefore
reduced the concentration of fu MO (300 μM), when injected into MZdhc2 mutants.
These injected MZdhc2 embryos failed to express nkx2.2 (Fig. 18B). Interestingly, the
expansion of the ventral intermediate gene, olig2, was also rescued (Fig. 18B). These
results demonstrate that fu is indispensable for Hh signaling in wild-type and mutant
medaka embryos and its overexpression augments the signal.
fu is known to be expressed ubiquitously in zebrafish at early developmental
stages (Xia et al., 2010), but the precise pattern and regulation of fu expression during
neural tube patterning have not been reported. My further analysis revealed that fu
expression is restricted to the ventral part of neural tube where high to low levels of Hh
31
signaling are activated at 16-somite stage in medaka (Fig. 19A). Furthermore, fu
expression was dorsally expanded in MZdhc2 neural tubes (Fig. 19A), like the ventral
intermediate genes. These results suggest that fu is a transcriptional target of Hh
signaling. To test this possibility, I treated wild-type embryos with 5 μM cyclopamine
and observed severe reduction or loss of fu expression in cyclopamine-treated embryos
(Fig. 19B), indicating that fu expression is induced by Hh signaling downstream of Smo.
I also confirmed that fu expression in zebrafish is ventrally restricted in the neural tube
and depends on Hh signaling (Fig. 19C).
I finally asked if Fu, when overexpressed, can restore Hh signaling, when
Smo-mediated signaling is compromised. For this, embryos were treated with 2.5 μM
cyclopamine (intermediate dose, Fig. 14A) together with fu mRNA injection. Those
injected embryos showed weak but significant up-regulation of nkx2.2 (n=12/18) as
compared with cyclopamine-treated control embryos (n=1/14) (Fig. 19D), suggesting
that Fu augments Hh activity downstream of Smo. Given that Fu is a positive mediator
of Hh signal transduction, Fu is likely to form a positive feedback loop downstream of
Smo to reinforce Hh signal in teleost target cells (Fig. 20).
32
Discussion
In the present study, utilizing the medaka mutant with severely shortened cilia, MZdhc2,
I demonstrated that shorter cilia mediate less Hh activation in fish, although the Hh
defects were milder than zebrafish mutants which completely lack cilia. This result
suggests that Hh activity in fish depends on the length of cilium. And I also found that
the receptor Ptch1 is localized on the cilium in fish. These are largely consistent with
the observation of murine ciliary mutants. Furthermore, the present study has addressed
why the expression of low-threshold target genes is expanded in ciliary mutant neural
tubes and how Hh signal is augmented in fish mutant cells.
A possible role of cilia in Hh gradient formation
olig2-positive wild-type cells in MZdhc2 neural tubes were positioned more dorsally
than dorsal boundary of olig2 expression in wild-type neural tubes (Fig. 17C). This
result, though indirectly, suggest that the gradient profile of Hh ligand dorsally shifts in
the mutant neural tubes. Additionally, consistent with this result, when I treated the
33
mutant embryos with various concentrations of cyclopamine, in the WT and Mdhc2
group, the percentage of olig2-positive embryos started to decrease at a cyclopamine
concentration as low as 2.5 μM, and went down below 27% at 5 μM, while 78% of
embryos maintained olig2 expression in the MZdhc2 groups (Fig. 14B). These results
indicate that in the case of the lower threshold olig2 expression, the activity of Hh
signaling in mutant cells is enhanced, probably because the gradient of Hh ligand was
dorsally expanded in the mutant embryos.
Dorsal expansion of Shh ligand was directly observed with smoothened mouse
mutants (Chamberlain et al., 2008) and this can be interpreted as a consequence of the
reduced amount of Ptch1 receptor, a downstream target of Hh signaling. Indeed, it has
been proposed that the Shh gradient is regulated by a Shh-induced negative-feedback
mechanism in which ligand binding to Ptch1 at the cilia sequesters Hh ligand itself in
the intercellular space (Jeong and McMahon, 2005). It is thus conceivable that in ciliary
mutant neural tubes, the reduced amount of Ptch1 on cilia caused a dorsally shifted Hh
gradient (Fig. 17C), and thereby the expression domain of ventral low-threshold target
genes is expanded, although further confirmation by direct imaging is required.
34
Consistently, the neural tube in Dnchc2 mutants also exhibits the expansion of low
threshold gene expression (Huangfu and Anderson, 2005; May et al., 2005). Thus in
vertebrates, the length of cilia (or the receptor Ptch1) could be one of the factors that
affect the Hh gradient in the neural tube.
Importance of cilia in Hedgehog signaling
In the present study, I generalized that cilium is required for Hh signaling in fish.
Additionally, the receptor Ptch1 was found to localize the cilium, as in mouse.
Considering the advantages of the use of cilia in Hh signal transduction, the
efficiency for signal transduction could be increased if cilia are centered. The length of
primary cilium is around 1-10 μm and the width is 500 nm. The volume of the cilia is
around 1/10000 of cytosol. They can thus concentrate signal-transducing molecules
10000 times higher if the molecules are only localized to the cilia; in the case of 1 μM
molecules localized to cilia, around 600 molecules are in the cilia, suggesting that the
efficiency for signal transduction could be greatly increased. Additionally, as described
in Introduction, cilia are thought to resemble the nucleus because of the high
35
concentration of GTP-bound Ran in the cilia and some nuclear pore proteins localized at
the ciliary base (Dishinger et al., 2010; Kee et al., 2012). Cells could use cilia to form a
compartment which is distinguished from the cytoplasm. In the case of Hh signaling,
the ciliary compartment is a good place to avoid mis-activation of the target genes
through ciliary sequestration of its transcription factor Gli. Additionally, considering
that a ciliary localization signal is known to be similar to a nuclear localization signal
(Dishinger et al., 2010), the translocation of ciliary proteins to the nucleus could be a
relatively easy strategy. Given that the use of the cilium has many advantages as
described above, fishes have evolved to use the cilium as a center of Hh signaling.
Furthermore, in addition to Ptch1, Smoothened in fish is known to localize to the cilium,
as in mammals (Aanstad et al., 2009). On the other hand, fishes maintain the
fused-mediated pathway which is lost in mammals. These facts highlight fish as a
unique model with which to investigate the transition state from non-cilia to
cilia-mediated Hh signaling.
36
Significance of teleost-specific augmentation of Hh signaling mediated by Fused
In the present study, I propose that Fu is a key component to account for the difference
in activation level between mammals and fish in ciliary mutants. Fused is a crucial
mediator of Hh signaling in Drosophila and zebrafish, but not in mammals (Wilson et
al., 2009). I first confirmed that fu is required for Hh signaling in medaka like zebrafish,
and next found that the expression of fu in the neural tube is restricted to the ventral part
and induced by Hh signaling in fish. Subsequent analyses demonstrated that Fu forms a
positive-feedback loop downstream of Smo (Fig. 20); Fu activates the Hh pathway
which then leads to the up-regulation of Fu. The positive feedback centered by Fu could
augment Hh signal in ciliary mutant cells with lower input of Smo-mediated signaling.
This could thus explain why the phenotype of fish ciliary mutants is milder than that of
mammalian counterparts. Of course, Fu may not be a mere component that
differentiates the ciliary dependency in the two vertebrate models. Indeed, in zebrafish,
low levels of Hh activation mediated by Gli1 is known to occur in an Hh-independent
manner and its mechanism remains elusive (Huang and Schier, 2009).
37
What is the biological and evolutionarily significance of the positive-feedback
mechanism in Hh signaling? A hint could be found in the speed and mode of neurulation
in fish. According to the recent report by Xiong et al. (2013), specification of neural cell
types in zebrafish begins earlier and proceeds faster under the noisy conditions of cell
movements in the formation of the neural keel, whereas in other vertebrates, such as
chick and mice, neurulation proceeds gradually and steadily in an epithelialized cell
sheet, following an established Shh gradient. A rapid and amplified response to Shh in
target cells would thus be necessary in fish neurulation.
Additionally, the size of neural tube in medaka is about one-fourth as large as
that in the mouse. To make Hh gradient correctly in the ventricular zone in medaka,
similar to that in mouse, it is likely that the amount of Hh ligands is lower in medaka,
suggesting the requirement of augmentation of Hh signaling in fish. Of course, the
quantification and/or the imaging of the ligand will be necessary to discuss this further.
Xiong et al. (2013) also showed that specified neural progenitors sort to form
sharply bordered domains from mixed progenitor populations. However, this apparently
contradicts the transplantation result showing ectopic expression of a specific marker in
38
wild-type donors (Fig. 17C), suggesting that multiple strategies, including sorting and
position-dependent determination, are employed to achieve a robust patterning.
Finally, the presence of cilium-mediated signaling was recently reported in the
olfactory epithelium of Drosophila (Kuzhandaivel et al., 2014), suggesting the
evolutionarily ancient origin of this mechanism. Thus, further analysis of Hh signaling
in diverse species and tissues will provide greater insight into the evolution of this
crucial signaling pathway.
39
Conclusions
The present study not only strengthens the idea of a conserved role of primary cilia in
Hh-signal transduction in vertebrates, but also uncovered a teleost-specific
augmentation mechanism mediated by Fu. The fish-specific augmentation can serve as
the mechanism that accounts for the lower cilia-dependency for Hh signaling in fish and
gives novel insight into the evolution of Hh signaling.
40
Materials and methods
Fish strains
All studies of medaka (Oryzias latipes) were carried out using d-rR strain of a closed
colony. ENU-based mutagenesis of medaka was performed as described previously
(Ishikawa, 1996). Zebrafish (Danio rerio) were Riken wild-type, strain RW. Fish stocks
were maintained under a long-day photoperiod of 13.5:10.5 h light:dark at 28 °C. Under
these conditions, medaka spawn daily within 1 hour of the onset of light for a number of
consecutive days, zebrafish spawn about once per two weeks. Collected medaka
embryos were sorted into medaka Hatching buffer, maintained at 32°C, and staged
according to morphological criteria (Iwamatsu, 2004). For zebrafish, the embryos were
sorted into 1/3 Ringer (39 mM NaCl, 0.97 mM KCl, 1.8 mM CaCl2, 1.7 mM HEPES at
pH 7.2) and maintained at 28.5°C and staged according to hours postfertilization (hpf)
at 28.5°C and morphological criteria (Kimmel et al., 1995). All experimental procedures
and animal care were carried out according to the animal ethics committee of the
University of Tokyo.
41
Genotyping of medaka aA90/dhc2 mutant
A small segment of caudal fin was excised and fix them in 100% MeOH. And genomic
DNA was extracted with 50 µl of DNA extraction buffer [10 mM Tris pH 8.0, 50 mM
KCl, 0.3% Tween20, 0.3% NP40 and 1 mg/ml proteinase K (Invitrogen)] at 55°C
overnight. The sample was then heated to 98°C for 10 min to inactivate proteinase K
and centrifuge at 15.3 kG, 4°C, 10 min and 2 µl of the supernatant of each sample was
used as a template for polymerase chain reaction, PCR (total volume 10 µl). PCR
primers were described in Table 2.
Antibody generation
His-tagged N-terminal (169-405; Ptch1-N-His) polypeptide of medaka Ptch1 (Fig. 16C)
was expressed in E. coli Rosetta (DE3) competent cells using pET24a (Novagen) vector.
Recombinant proteins were purified with Profinity™ IMAC Ni-charged resin (Bio-Rad)
under denaturing conditions and dialyzed against PBS. This Ptch1-N-His polypeptide
was used for immunization of rabbits. Anti-medaka Ptch1-N-His (169-405) antibody
42
was affinity-purified using Immobilon membrane (Millipore) strips onto which the
antigen were transferred as described (Okano-Uchida et al., 2003).
Whole mount in situ hybridization
Embryos were fixed with 4% paraformaldehyde (PFA) in PBST (phosphate-buffered
saline containing 0.1% Tween-20) overnight and dechorionated manually with forceps,
then stored in MeOH at –20°C. After rehydration, embryos were permeabilized with
proteinase K (10 μg/ml) at room temperature for 7 minutes and re-fixed with 4%
PFA/PBST for 25 min. The specimens are washed five times in PBST. Hybridization
was carried out at 65°C with digoxigenin-labeled probes overnight. Hybridized embryos
were washed with 50% formamide/2xsaline-sodium citrate (SSC)-0.1% Tween20
(SSCT), 2xSSCT and 0.2xSSCT, incubated in 0.2% blocking reagent (Roche) at room
temperature for more than 2 hours, and then treated with anti-digoxigenin antibodies
labeled with alkaline phosphatase (Roche) at 1:7000 dilution with 0.2% blocking
reagent. The staining reaction was started by incubating medaka embryos with
NBT/BCIP solution according to the protocol described for the NBT/BCIP ready-to-use
43
tablets (Roche), zebrafish embryos with BM Purple (Roche). The cDNAs used as the
templates for the probes were described in Table 2.
mRNA overexpression
mRNAs were synthesized as reported previously (Yokoi et al., 2007). Briefly, to
synthesize RNA in vitro, the open reading frame of the medaka ptch1 and fused were
cloned into the pCS2+ vector. The dominant-negative regulatory subunit (R1α) for
murine PKA (dnPKA) was kindly provided by Dr. Anna Wild (Univ. of Sheffield).
Capped sense RNA was synthesized using mMESSAGE mMACHINE® SP6
Transcription Kit (Ambion). The synthesized RNA was purified using the RNeasy Mini
kit (QIAGEN) and stored at −80°C until use.
Microinjection and Cell transplantation
To generate the maternal-zygotic aA90/dhc2 mutant, donor embryos were obtained from
intercrosses of aA90/dhc2 heterozygous fish; host embryos were from olvas-GFP
transgenic fish, whose oocytes were labeled with green fluorescent proteins using the
44
regulatory region of the medaka vasa gene (olvas, named after Oryzias latipes vasa)
(Tanaka et al., 2001). Donor embryos were injected at one-cell stage with 10 mg/ml
rhodamine-dextran (10 kDa; Molecular Probes). Host embryos were injected at one-cell
stage with 300 μM of a morpholino antisense oligonucleotide directed against dead end
mRNA. At morula stage, donor and host embryos were dechorionated with hatching
liquid (Yasumasu et al., 1989). At mid-blastula stage, the embryos were placed on
V-shaped grooves of a 1.5% agarose gel immersed in Yamamoto’s Ringer (Yamamoto,
1956), and then the 50–100 cells were transplanted from the margin of donor embryos
into the animal pole of similarly staged hosts using a micromanipulator (M-152,
Narishige) in combination with a microinjector (IM-6, Narishige). Transplantation
needles were made from a glass capillary (G-1, Narishige) pulled by a horizontal pipette
puller, and clipped with forceps to form a sharp tip. Donors and hosts were cultured
individually in glass dishes until 2 dpf; host embryos were then screened for the transfer
of fluorescently labeled donor-derived PGCs, and donor embryos were genotyped for
dhc2. To confirm the PGC elimination in host germ cells, these host embryos were
45
checked whether GFP signals were disappeared at hatching stage. If germ cells
elimination were not succeeded, the offsprings will have GFP fluorescent signals.
For normal cell transplantation, donor embryos were injected with the mixture
of rhodamine- and biotin-dextran. At mid-blastula stage, scores of cells were
transplanted into the margin of donor embryos to make mosaic neural tube.
Histology
The paraformaldehyde-fixed fish embedded with 1.5% Agarose in 1/2 hatching buffer
were dehydrated in a graded series of ethanol-water mixtures, then incubated in 1:1
ethanol and infiltration solution (Technovit 7100, Heraeus Kulzer) for 30 min. The
specimens were incubated in infiltration solution for at least 4 hours and positioned in
polymerization medium (Technovit 7100, Heraeus Kulzer) overnight at 4°C and
sectioned with RM2245 (Leica) at a 7-12 µm thickness. Microtome sections were
prepared and mounted on slides, air dried at 30°C, stained with Nuclear fast red (Vector)
or Mayer's Hematoxylin Solution, overlaid with rapid mounting medium (Entellan new,
Merck), and cover-slipped.
46
Immunohistochemistry
For immunohistochemistry, medaka embryos were dechorionated using hatching liquid
in 1/2 Yamamoto Ringer, fixed with 4% PFA/PBS for 1 hour at room temperature,
permeabilized with 0.5% Triton X-100 in PBS at room temperature for 30 min, rinsed
with PBS, incubated in blocking solution (2% BSA, 10% DMSO, and 0.2% Triton
X-100 in PBS) for 1 hour at room temperature, and then incubated with primary
antibodies in blocking solution overnight at 4°C: Ptch1 at 1:200; Monoclonal
anti-acetylated α-tubulin antibody (T6793, Sigma) at 1:400; monoclonal anti-γ-tubulin
antibody (T3559, Sigma) at 1:1000. After washing in PBSDT (1% DMSO and 0.1%
Triton X-100 in PBS), embryos were incubated with Alexa 488, 555 or 647 (Molecular
Probes) conjugated secondary antibodies at 1:400 in blocking solution overnight at 4°C.
Embryos were washed with PBSDT at room temperature after incubation with each
antibody. Washed embryos were cleared in 50% glycerol/PBS and photographed on a
LSM710 confocal fluorescence microscope (Zeiss). For the cilia staining of neural tube,
47
the apical surface were exteriorized with forceps prior to fixation. Next steps are
according to the normal immunohistochemistry protocol.
Chemical treatment
The chorion was manually removed using hatching liquid for medaka and 5 mg/ml
pronase for zebrafish, prior to the incubation. To inhibit Hh signaling dechorionated
embryos were incubated from 30-50% epiboly stage in cyclopamine (BML-GR334,
Enzo Life Sciences). Control embryos were treated simultaneously with an equal
concentration of DMSO.
RT-PCR
RT-PCR analysis was performed using total RNA of medaka embryos. Total RNA was
isolated from medaka embryos at the late gastrula stage using ISOGEN (Nippon gene),
according to the manufacturer’s instructions. First-strand cDNA was prepared from 5 µg
total RNA using an oligo d(T) primer and SuperscriptIII reverse transcriptase
48
(Invitrogen) and cDNA fragments were obtained by PCR with the primers described in
Table 2.
PCR fragments were then cloned into the pCRII-TOPO vector (Invitrogen). For
the RNA probe synthesis, the insert was transcribed in vitro with SP6 or T7 RNA
polymerase (Promega) after linearization of the plasmid.
Scanning electron microscope
Medaka embryos were dechorionated using hatching liquid in 1/2×Yamamoto ringer.
Apical surfaces of neural tube were exteriorized with the forceps. These embryos were
immediately fixed in 2.5% glutaraldehyde (TAAB), 2% paraformaldehyde (Thermo) in
0.1 M sodium phosphate buffer pH 7.4 (Wako) for 18 hours at 4°C. These embryos
were postfixed in a 1% osmium tetroxide solution (TAAB) for 2 hours at 4°C,
dehydrated in an ascending series of ethanol and infiltrated with t-butyl alcohol
(Nacalai). The specimens were frozen at 4°C, freeze-dried, mounted on aluminums tubs
with double-sided adhesive tape, coated with platinum, and viewed at 10 kV on a
Hitachi S-800 scanning electron microscope.
49
Figures
50
Figure 1. The French flag model providing a positional information by a
morphogen concentration gradient.
In this model, the positional information is delivered by a gradient of a diffused
morphogen extended from a source cell (green). Morphogen forms a concentration
gradient at the extracellular space. Multiple cell types are differentiated in a
concentration dependent manner. Figure based on Rogers & Schier, 2011.
51
Figure 2. Hedgehog signal transduction pathway.
(A) In the absence of the Hedgehog (Hh) ligand, the downstream transcription factor
Gli/Ci is cleaved to its repressor form through PKA-dependent phosphorylation. (B)
When Hh ligand binds to Patched (Ptch), the conformation of Ptch is altered, releasing
the inhibition of Smoothened (Smo). Smo then inactivates PKA, and Gli/Ci is
translocated into the nucleus and activates the transcription of Hh target genes.
52
Figure 3. Schematic view of neural tube patterning by Shh concentration gradient.
In the neural tube, Sonic hedgehog (Shh) ligand forms a dorso-ventral gradient from its
source, notochord and floorplate. The specific genes for the progenitor domains are
expressed in a Shh-ligand concentration dependent manner.
53
Figure 4. The formation and maintenance of cilia mediated by the intraflagellar
transport.
A cilium is a microtubule-based organelle, arise from a basal body anchored to the base
of cilia by transition fibers. Ciliary proteins and membrane receptors are delivered by an
intraflagellar transport machinery along the ciliary axoneme (microtubule). The
anterograde transport, from the basal to tip, is powered by the kinesin, and the
retrograde transport, from the tip to the basal, by the cytoplasmic dynein. Most of cilia
54
have a ciliary pocket at the base, distinguished from the microvilli, which is an
actin-based cellular projection and narrower than the cilium.
55
Figure 5. Morphological phenotypes of aA90/dhc2 mutants.
(A-D) Frontal views of the heart at 6 days post fertilization (dpf). In wild type (WT)
embryos, the ventricle of the heart positions to their right side and the atrium to the left.
Conversely, in zygotic mutants of dhc2 (Zdhc2) and maternal-zygotic mutants of dhc2
(MZdhc2), the ventricle of the heart positions to their left and the atrium to the right.
(E-L) Lateral views of the ventricle of the brain (E-H) and the somite (I-L) at 3 dpf.
MZdhc2 showed enlarged brain ventricles (H) and U-shaped somites (L) instead of
chevron-shaped ones (I-K). (M-T) Transverse section of nephric duct (M-P) and tail
56
morphology at 7 dpf (Q-T). MZdhc2 embryos exhibited expanded nephric ducts (P).
Severe ventral curvature was observed in MZdhc2 (T), compared with its zygotic
mutants (R). v, ventricle; a, atrium. Scale bars: 100 μm in D, H, L, P; 200 μm in T.
57
Figure 6. The medaka aA90/dhc2 lacks essential domains of the dhc2 gene.
(A) Positional cloning of the aA90 mutation in linkage group (LG) 13. The number of
recombinants at each marker is shown. aA90/dhc2 has a 37.7 kb deletion in the dhc2
locus including the start codon. Colored arrows (blue, red and green) indicates the
position of primers used for genotyping described in Fig. 6B.
(B) Genotyping for aA90/dhc2. The primers for PCR detection were described in arrows
of Fig. 6A and listed in Table 2. In aA90/dhc2 mutants, the aberrant product (+/- and -/-,
top panel) and the loss of the product (-/-, middle panel) were detected by genomic
PCR.
58
(C) Schematic diagrams of Dhc2 protein expressed in wild-type and the deleted region
in mutants. The mutant lacks the heavy chain (HC)-HC and the HC-Intermediate chain
interaction domain and the AAA ATPase domain of Dhc2. Colored arrows indicates the
primer set for the analysis of dhc2 expression, described in Fig. 6D. HC, heavy chain;
IC, intermediate chain; MT, microtubule; a.a., amino acids.
(D) Expression analysis of dhc2 at 6 dpf by RT-PCR using primers described in Fig. 6C
(arrows) and listed in Table 2. The dhc2 expression was diminished in homozygous
aA90/dhc2 mutants (-/-).
59
Figure 7. Generation of Maternal-Zygotic dhc2 mutant (MZdhc2).
Germ-line replacement strategy using the rhodamine-dextran labeling technique. (A)
Overview of transplantation strategy showing the transfer of cells from the margin of
rhodamine-dextran-labeled mutant donor embryos into the animal pole of dead end-MO
injected WT hosts (Tg[olvas-GFP], germ cells are labeled with GFP (Tanaka et al.,
2001)). Donor embryos are genotyped after transplantation. A morpholino antisense
oligonucleotide (Genetools) to dead end was complementary to a region covering the
splicing site for exon 2 and intron 2, 5'-TGTTCAGAACTGGCCTCTCACCATC-3'.
60
(B) Chimeric host embryos were screened at 2 dpf for the presence of
rhodamine-labeled donor PGCs that had migrated successfully into the gonadal
mesoderm (arrowhead). Host embryos also showed somatic contribution of
rhodamine-dextran-labeled donor cells to anterior neuroectoderm lineages (*). (C)
Chimeric host embryos were screened again at 4-6 dpf for the lost of GFP-labeled host
PGCs at the dorsal region of the gut (arrowhead). Scale bars: 500 μm in B-C.
61
Figure 8. Cilia are shortened in MZdhc2.
(A) SEM analysis of the ventricular surface of the neural tube at 16-somite stage. Cilia
were found on the surface of neural epithelial cells (arrowheads). In wild type and
Mdhc2, longer cilia were observed on the floorplate (VM, ventral midline), compared
with those on the lateral neural tube (LNT). Cilia in MZdhc2 were greatly shortened on
both regions. (B) Schematic view of opening of the apical surface of neural tube with
forceps. (C) Medaka MZdhc2 mutants have shortened cilia in Kupffer’s vesicle. Cilia
62
were visualized by staining with anti-acetylated α-tubulin antibody (green) and basal
bodies were visualized with anti-γ-tubulin antibody (magenta). Scale bars: 5 μm.
63
Figure 9. Neural tube patterning in MZdhc2 mutants.
(A) Expression of neural tube markers in a cross-sectional view at 16-somite stage (The
dashed line in Fig. 10A indicates the section plane). All Hh target genes were expressed
in MZdhc2, but olig2 and nkx6.1/6.2 expression was dorsally expanded. The lower
64
panels of nkx2.2 are the magnified images of the upper panels (dotted line). (B)
Representation of the size of each progenitor domain along the DV axis in WT, Mdhc2
and MZdhc2. Scale bar represents 20 μm.
65
Figure 10. Gross patterning of neural tube and somite in MZdhc2 mutant.
(A-J) Expression of neural tube markers in MZdhc2 mutant embryos. Wild type, Mdhc2
control medaka embryos and MZdhc2 mutants were stained at 16-somite stage for the
66
expression of dbx2 (A), dbx1 (B), pax3 (C), pax6 (D), nkx6.2 (E), nkx6.1 (F), olig2 (G),
nkx2.2 (H), foxa2 (I), shh (J), shown in a lateral view. MZdhc2 mutants show shh, foxa2
and nkx2.2 expression (H-J), dorsally expanded expression of olig2, nkx6.1 and nkx6.2
(E-G; arrowheads), and retracted expression of dbx genes, pax6 and pax3 (A-D;
arrowheads), compared with those expression in the control embryos. * in D indicates
somites. Cross-sectional views at the dashed line in A were depicted in Fig. 9A. (K)
Somite patterning in MZdhc2 embryos. Adaxial cells (engrailed1-positive cells) were
significantly decreased in MZdhc2 as compared with control embryos. (L) ptch1
expression in MZdhc2 is nearly identical to that in WT. Scale bar: 500 μm.
67
Figure 11. Dose-dependent effects of cyclopamine treatment on the expression of
Hh target genes.
(A) foxa2, nkx2.2 and olig2 expression in WT embryos treated with DMSO, 2.5 μM and
5 μM cyclopamine (cyclop). (B) foxa2 expression is absent in the lateral FP and only
detectable in the medial FP in embryos treated with 5 μM cyclopamine, when compared
to the DMSO control. Scale bars: 100 μm in A, 20 μm in B.
68
Figure 12. nkx2.2 and olig2 expression at three anterior-posterior axis levels.
(A) nkx2.2 and olig2 expression at three different anterior-posterior axis levels of
16-somite stage embryos. (B) olig2 expression in WT for indicating the position of
anterior (A), middle (M) and posterior (P) level, depicted in A and C. (C) Measurement
of the dorsal boundary of nkx2.2 and olig2 expression at relative distances (percentage
69
(%) of the neural tube) from the floor plate in WT and MZdhc2 of 16-somite stage (n ≥
3 embryos; mean ± SD). For the representation of the dorsal boundary of nkx2.2 and
olig2 expression in same graph, blue shade is for nkx2.2 and red shade is for olig2
expression. The olig2 boundary in mutant embryos is significantly different from WT
counterparts (p values from Student's t test: Anterior, p < 0.05; Middle, p < 0.0005;
Posterior, p < 0.005). Scale bar represents 20 μm.
70
Figure 13. A schematic drawing explaining the similarities and differences in
ciliary and neural tube phenotypes between fish and mouse dhc2/dnchc2 mutants.
Ciliary phenotypes and dorsal expansion of olig2 domain in MZdhc2 mutants are nearly
identical to those in mouse mutant, but nkx2.2 expression was reported to be lost in
mouse mutant (Huangfu and Anderson, 2005; May et al., 2005). D, dorsal; V, ventral.
71
Figure 14. Hh signaling activity is partially defective in MZdhc2 mutants.
(A-B) The percentage of nkx2.2 (A), olig2 (B) positive embryos with the graded series
of cyclopamine treatment (Sample numbers are indicated in Table 4). (C) Dorsal view
of nkx2.2 expression in 0.5 μM cyclopamine treated and control (DMSO-treated)
embryos. Scale bars: 100 μm in C.
72
Figure 15. MZdhc2 is sensitive to dnPKA.
dnPKA mRNA injected MZdhc2 exhibited expanded nkx2.2 expression (n=19/24,
arrowhead), consistent with dnPKA mRNA injected-WT (n=22/24, arrowhead) and
Mdhc2 embryos (n=17/20, arrowhead), compared with Control (WT) embryos. Scale
bar: 500 μm
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74
Figure 16. Ptch1 is localized to the cilia in medaka fish.
(A) Phylogenetic trees showing the relationship between Patched proteins across
vertebrates based on neighbor-joining method using ClustalX and maximum likelihood
estimation using RAxML. o, Oryzias latipes (medaka); g, Gallus gallus; h, Homo
sapiens; m, Mus musculus; x, Xenopus tropicalis; d, Danio rerio. In the both methods,
medaka Ptch1 is the homologue of mammalian Ptch1, not Ptch2. All sequences are
obtained from Ensembl Web site and the accession numbers are listed on Table 3. (B)
Injection of ptch1-morpholino antisense oligo for splicing blocking (intron 5 and exon
6) (5'-CCCCTACCTCTGTAAAGTTAATTAC-3') induced ectopic Hh-dependent
muscle pioneer (Eng+ cells, lateral view, arrowheads), visualized by staining with
anti-Engrailed antibody (4D9). (C) The recombinant protein of His-tagged N-terminal
(169-405; Ptch1- His) polypeptides of medaka Ptch1 (orange lined) were expressed
using pET24a vector and the polypeptides were used for immunization of rabbits. (D)
Ptch1 were visualized by staining with anti-medaka Ptch1 antibody (green) and cilia
were visualized with anti-acetylated α-tubulin antibody (red). Ptch1 morpholino oligo
injected embryos (ptch1 MO) had no Ptch1 positive signals at 16-somite stage. (E)
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Ptch1-myc (magenta) was specifically localized to cilia in neural tube and the signals
are well merged with anti-Ptch1 antibody signals (green) in a cross-sectional view at
16-somite stage (arrowheads). The inset on the right side is a high magnified image of
the square region. Scale bars: 50 μm in B, 5 μm in D, 10 μm in E.
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Figure 17. Ectopic olig2 expression of WT cells in the dorsal region of MZdhc2
neural tube.
(A) Schematic view of the transplantation of biotin-injected WT cells (brown colored)
into Mdhc2 (control) and MZdhc2 embryos. (B) Dorsal views of the eyes in WT, Zdhc2,
Mdhc2 and MZdhc2 at 16-somite stage. Optic cup and lens formation were significantly
defected in MZdhc2, as compared with WT, Zdhc2 and Mdhc2. Arrows indicate optic
cup and arrowheads indicate lens. (C) olig2 expression in the transplanted embryos.
Ectopic olig2 expression (purple) of WT cells (brown) was observed in the dorsal
77
region of MZdhc2 neural tube (arrowhead, right panel), not in that of Mdhc2 one
(arrowhead, left panel). Scale bars: 100 μm in B, 20 μm in C.
78
79
Figure 18. fused is required for Hh signaling in medaka fish.
(A) Knockdown of fu was performed using the morpholino-oligonucleotide (MO) for
splice blocking (5'-CAACCACCTTATTGACGACAAAACA-3'). Diagram of altered fu
splicing in morphants of fu-i1e2 inserts intron 1 (+In. 1), resulting in an out-of-frame
truncation of the fu protein, and splices exon 2 to a cryptic acceptor in exon 3 (- Ex. 2),
causing an out-frame mutation of fu. The effect of the splice-blocking MO was verified
by RT-PCR from 20 embryos total RNA (16-somite stage). The positions of primers for
checking the effect of MO were indicated in A (arrows) and listed in Table 2. MO
caused splice-blocking effectively. (B) nkx2.2, olig2 and shh expression in fu-MO
injected embryos. nkx2.2 expression was almost diminished in 600 μM fu-MO injected
WT embryos (arrowhead) and greatly reduced in 300 μM fu-MO injected MZdhc2
embryos (arrowhead). The dorsal expanded expression of olig2 was rescued in 300 μM
fu-MO injected MZdhc2 embryos (arrowhead). (C) fu mRNA injection rescued nkx2.2
expression in fu-MO injected embryos. (D) fu overexpression induced ectopic nkx6.1
and olig2 expression (arrowheads) in a cross-section view and a lateral view (the dashed
line indicates the section plane).
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Figure 19. fused expression pattern in medaka and zebrafish, and fused augments
Hh signaling in medaka.
(A) fu expression in a cross-sectional view and a lateral view (the dashed line indicates
the section plane). fused expression was ventrally restricted. Dorsally expanded
expression of fu was observed in MZdhc2 (arrowhead). (B) fu expression (arrowhead)
in the neural tube was lost in 5 μM cyclopamine-treated embryos. Black lines mark the
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magnified areas, depicted in the right panel. (C) fu expressed in a Hedgehog-dependent
fashion also in zebrafish. The embryos treated with 100 μM cyclopamine did not
express fused or nkx2.2a (a Hh target gene). Arrowheads indicate the expression in the
neural tube. The dashed line indicates the section plane. (D) The loss of nkx2.2
expression in 2.5 μM cyclopamine-treated embryos (n=13/14) was rescued by
overexpression of fused (n=12/18, arrowhead). Scale bars: 500 μm in A (lower panel), B,
C (left panel), D; 20 μm in A (upper panel), C (right panel).
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Figure 20. Proposed model of the distinct features of Hh signal transduction in
insect, fish and mammal.
fu is expressed in a Hedgehog-dependent fashion and is also one of the components of
the Hedgehog pathway in fish. Fused negatively regulates Suppressor of Fused (SuFu),
which is a negative regulator of Gli/Ci in Hh signaling. The transcription of fused in fish
could lead to Hh activation. This positive-feedback loop amplifies Hedgehog pathway
in fish downstream of cilia.
83
Tables
84
Table 1. Defects in heart asymmetry in dhc2 mutant embryos and morphants
Genotype n Correct (%) Reversed (%)
Wild type 110 99.1 0.9
Zdhc2 294 76.2 23.8
dhc2 MO* 108 77.8 22.2
Mdhc2 82 100 0
MZdhc2 128 46.9 53.1
* dhc2-Met MO, 5'-AAATGCGGCAGACTCGCAGTTTTAC-3’
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Table 2. Primers used in this study
Genotype Forward (5' to 3') Reverse (5' to 3')
primers I CCCGCTGAGTTTAGAGACTATTG GTGGGAAGTGTACACCTTCATAAT
primers II TTCTATGGGTGATGCCACTTTC GGAAATCTGATACAACCCCAGC
primers III CTCAAAGTGAGCTTTTGGCTCAAGTATT ACTGTAGAAGATGGGACACGAAGAAAAG
Probe* Forward (5' to 3') Reverse (5' to 3')
nkx2.2 TCGTTGACCAACACAAAGACGG CCAAGTCCTGAGCTTTAAGAGTGTG
olig2 ATACAAGTCGTGTGTCAAGCAGACC TGAGAAGTCCGTGATGGGGTC
fused TTCAGTAAAAACGCGTGAGC AACACGTTTGTGTCCGACAG
nkx6.1 TCTTCTGGCCGGGAGTCATG AAGTGCTTTACATGAAGCTGCG
nkx6.2 ATGGAAGCTAACCGGCAGAG CACTTGGTCCTCCGGTTCTG
pax3 CAGGAGGTTTACCAAGAATGATG AAGACTGAGTACTGGGCAGAGTG
dbx1 AAGAAGCGGTTCCTGATTTCTC CTCATTCTTTCTCCTCCCAACTC
dbx2 CTCCTGCTCTGCCAGGTTTTG CACTGGTGTGATTGTGTGACAG
eng1 AACCACCAACTTTTTCATCGAC ATCTGGGACTCGTTCAGGTG
zebrafish nkx2.2a GCACTCCTTACTTTCATTTGG CGTATAACACGAAGGACAAAAG
zebrafish fused GGAGAAAACGGTCTAAGTTATG ATCAGAACTCCATCTGCAAC
*shh (AB007129) and foxa2 (AB001572) were kindly provided by Dr. K. Araki; pax6 were by Dr. A. Kawakami.
RT-PCR Forward (5' to 3') Reverse (5' to 3')
fused ATGAATTCCTATCACGTCTTG ATGCAGTTATCACTCATTGTGTC
β-actin GATGAAGCCCAGAGCAAGAG AGGAAGGAAGGCTGGAAGAG
dhc2 (ex40-45) GTGCAAGCACTGAGGCTC CACTAGACTAGTTTCCACCACAAAG
dhc2 (ex86-98) CTTTGTCCACGGCCTGTTC CTGTTTGAGAAAAAGAGCAGCTC
dhc2 (del) GTTGAGGTGTGGTTAGGAGAGC TGGTTTGCTCATGGCTACG
86
Table 3. Accession numbers used to create the phylogenetic trees depicted in Fig.
16A.
Ptch1 Ptch2
Oryzias latipes ENSORLG00000004345 ENSORLG00000016137
Gallus gallus ENSGALG00000012620 ENSGALG00000010133
Homo sapiens ENSG00000185920 ENSG00000117425
Mus musculus ENSMUSG00000021466 ENSMUSG00000028681
Xenopus tropicalis ENSXETG00000014834 ENSXETG00000018892
Danio rerio ENSDARG00000016404 ENSDARG00000055026
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Table 4. Number of samples to examine Hh activity with the graded series of
cyclopamine treatment depicted in Fig. 14A-B.
cyclopamine (nM) 250 500 1000 2500 5000
nkx2.2 Wild type 20 23 26 18 17
Mdhc2 18 19 20 20 18
MZdhc2 20 26 19 18 16
olig2 Wild type 17 22 19 26 26
Mdhc2 16 22 18 18 14
MZdhc2 15 12 13 19 18
88
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Acknowledgements
I am using this opportunity to express my gratitude to everyone who supported me
throughout the course of my research.
First and foremost, I would like to express my deepest appreciation to my
supervisor, Dr. Hiroyuki Takeda for his substantial support, encouragement and patience.
My sincere gratitude is reserved for the other members of my committee, Drs.
Yoshitaka Oka, Masanori Taira, Manabu Yoshida and Mariko Kondo for their
constructive advices and valuable comments to my doctoral thesis.
I would also like to express my special appreciation and thanks to Dr. Haruo
Hagiwara for his experimental support on SEM analysis; to Dr. Keiichiro Kamura, Dr.
Tadashi Ishiguro and Mr. Yohei Masuda for their contributions to the identification of
aA90 responsible gene and their help in starting this work; to Drs. Sumito Koshida,
Tatsuya Tsukahara, Atsuko Shimada and other members of Takeda Laboratory for their
helpful discussions and suggestions; to Ms. Yasuko Ozawa for her excellent fish care.
93
Finally, I am greatly indebted to my family, Kayoko, Uichiro and Fumiko
Yamamoto. Words cannot express how grateful I am for their heartfelt support,
encouragement, suggestions and unconditional affection without which I would not
have accomplished this study.